ATR kinase supports normal proliferation in the early S phase by preventing replication resource exhaustion

The ATR kinase, which coordinates cellular responses to DNA replication stress, is also essential for the proliferation of normal unstressed cells. Although its role in the replication stress response is well defined, the mechanisms by which ATR supports normal cell proliferation remain elusive. Here, we show that ATR is dispensable for the viability of G0-arrested naïve B cells. However, upon cytokine-induced proliferation, Atr-deficient B cells initiate DNA replication efficiently, but by mid-S phase they display dNTP depletion, fork stalling, and replication failure. Nonetheless, productive DNA replication and dNTP levels can be restored in Atr-deficient cells by suppressing origin firing, such as partial inhibition of CDC7 and CDK1 kinase activities. Together, these findings indicate that ATR supports the proliferation of normal unstressed cells by tempering the pace of origin firing during the early S phase to avoid exhaustion of dNTPs and importantly also other replication factors.

The manuscript "ATR kinase supports normal proliferation in the early S phase by preventing replication resource exhaustion" by Menolfi et al. explores the role of ATR in DNA replication on unperturbed B cells. The authors take advantage of synchronous cell cycle entry of B cell upon cytokine stimulation to knockout ATR in non-dividing cells and observe the effects of the knockout on DNA replication upon stimulation. While the essential function of ATR in cell cycle has been extensively studied before, using ATR inhibitors and cancer cells, the authors of the current manuscript take it one step further, using primary cultures and a genetic approach. The main conclusion of the current study is that ATR is required to suppress excessive origin firing during DNA replication to prevent nucleotide depletion and subsequent DNA damage. This has been documented in cultured cancer cells before, so the main novelty is demonstrating that the same principles apply in primary B cells. The question why nucleotide complementation failed to restore S-phase progression remains unanswered. While the manuscript contains a vast amount of data and most of the conclusions are justified, the conclusions are mainly confirming the results that were previously reported in cancer cells/cell lines. In order to make it more novel, I propose either looking into some unsolved questions related to the effect of ATR inactivation (insufficiency of nucleotide complementation, effects on the nucleotide balance, etc.), or focusing on possible differences between healthy B cells and cancer.
Additional specific concerns/suggestions: 1. ATR being most important in early S-phase was previously proposed by Zou lab (Buisson et al, 2015). Here the authors conclude that ATR is mainly important for early S-phase cells based on the experiment where ATRi was added at different timepoints during cell cycle progression (figure 2f). On this figure the authors compare 2h ATRi incubation with 48h ATR incubation. Given that the cells enter S-phase less that 24h after activation, the 48h sample has been lacking ATR activity through the whole cell cycle, while 2h sample has been without ATR activity for only 2h. Similarly, on figure S3c, the authors compare 24h or 14h ATRi treatment to 2h. ATRi treatment in early S should be limited to 2h as well -my guess is it would not completely stop EdU incorporation. 2. A better experiment is presented on Fig. 3e, where authors look at the ability of the cells to complete the cell cycle by quantifying the G1 phase cells (this could be repeated and statistics could be presented). However, here we can see that at 24h timepoint when the BrdU pulse is performed, much fewer cells in the ATRi samples are actually incorporating BrdU (13.5% vs. 36%), so this must be taken into account when quantifying the percent of BrdU+ G1 cells at 48h -the percent cells labeled with BrdU at 24h cells making it to G1 (and not the percent of all cells ending up BrdU+ in G1) may actually be higher in the samples with continuous 48h ATRi treatment (possible re-activation of CHK1 by DNAPK proposed by Buisson et al., 2015?). Additionally, the percent of BrdU+ cells appears to increase between 24h and 48h, which is difficult to explain given a short 30 min pulse. 3. Metabolome profile on figure 5d is remarkable. It is showing lower concentrations of the majority of nucleotides in ATR-deficient cells, however, ATP, GTP, UTP, CTP, and dGTP appear to have increased with the loss of ATR, but it not discussed at all. These data indicate specific biosynthetic pathways may be regulated by ATR. A recent preprint from Bakkenist lab (Sugitani et all, 2022) showed regulation of RRM2 by ATR in dividing primary T cells, could a similar mechanism explain this phenotype in B cells? 4. Throughout the manuscript, the authors compare their findings in B cells to the studies in yeast, yet they fail to reference some key studies done in mammalian cells -nucleotide complementation has been done with CHK1 depletion by Gottifredi lab (González Besteiro et al, 2019); CDC7 inhibition has been tried in human cells by Helleday lab (Petermann et al. 2010) with CHK1 inhibitors, the role of CDK1 in ATRi-induced effects has been described by Bakkenist lab . 5. Specifically, unable to fully complement the effect of CHK1 depletion with nucleotides, González Besteiro et al., 2019 concluded that CHK1 depletion creates replication barriers slowing down replication. It would be very interesting to see if similar barriers can be observed in primary cells lacking ATR.
Minor suggestions: 1. Figures 2a and f have BrdU/PI plots in an unusual orientation. They would be easier to view if PI was on the horizontal axis and BrdU on the vertical, as it is the standard in the field, and as the authors show it on figure 3, for example.
Reviewer #2 (Remarks to the Author): The study by Menolfi et al. entitled "ATR kinase supports normal proliferation in the early S phase by preventing replication resource exhaustion" aims at understanding the role of ATR in the proliferation of unstressed proliferative B-cells. They initially show that the genetic deletion of ATR, or the loss of its kinase activity, impairs clonal expansion in-vivo, but has no effect on G0-arrested naïve B-cells. To better understand the molecular mechanism by which ATR promotes B cells proliferation, they use a model of naïve B cells ex-vivo culture, stimulated by cytokines (mimicking B-cells activation) and used state-of-the-art techniques such as metabolomics and CRISPR-Cas9 screens, to uncover the determinants of ATR-dependent proliferation. The authors show that cells depleted of ATR, show defects in S-phase progression, and propose that this is due to the function of ATR in fine-tuning origin firing to limit dormant origin firing and prevent a shortage of replication factors, and/or dNTPs. The manuscript tackles an appealing aspect of ATR functions in normal proliferation and uses a relevant physiological model of B-cell activation. These observations are potentially interesting, but the novelty is limited by the fact that the role of ATR and CHK1 in controlling origin firing locally and globally, has already been well established in the context of replicative stress. The study suffers a lack of evidence supporting their views and few technical limitations. Therefore, I do not think that this manuscript is suitable for publication in Nature Communication in its current form.
Major points 1-The authors show that CDK1/CDC7 dual inhibition restores the proliferation of ATR deficient cells. This is a critical result that has potential applications in several pathologies. However, the relevance of these findings needs to be confirmed in an in-vivo setting, such as in the context of naïve B cell clonal expansion 2-Linked to major point 1, do patients with Seckel syndrome suffer from a deficit of naïve/activated B-cells? Is it recapitulated in the mouse model of Seckel syndrome?
3-The authors should show representative images of the DNA fibers, and estimate the inter-origin distance, or the percentage of first-labels origins to support their findings on origin firing. 4-Analysis of cell viability by SSC/FSC gating is not accurate and can be misleading. The authors should confirm those results with a viability dye. 5-In Supplementary Figure 2d, Hydroxyurea did not affect S-phase progression? Was it used at a suboptimal concentration? The authors should confirm that HU and the ATM/DANPKcsi inhibitors hit their targets in this cellular model. Figure 2E, the authors should show the expression of ATR by Western-Blot 7-Can the authors provide more information about the screens, e.g. the raw and adjusted p values of the individual gRNAs? Why use an FDR<0.2 and not the more classically used FDR<0.1 or FDR<0.05? 8-The findings from the screens are very interesting but I see a biological bias that the authors did not comment on. How many of these gRNAs totally prevent origin firing? In this situation, they would confer resistance to ATRi, not because they are ATR targets but because they impede S-phase entry. For example, I would expect to find gRNAs of genes that induce G1-arrest (Mdm2, p21 ubiquitin ligases,…). Comparing gRNA representation before and after the 6days treatment may clarify those points.

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Minor points 1-The authors did not mention the gating strategies used in figure 1a before the B220/CD43 and B220/IgM plots. Did they exclude dead cells (e.g. 7AAC-or DAPI-cells)?
2-The statistical analysis is missing in figure 1b. Figure 7A, the legend is missing 4-the authors show that ATR depletion induces DNA damage in quiescent B-cells. This is rather surprising. Can the authors comment?

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Reviewer #3 (Remarks to the Author): In this manuscript, Zha and colleagues examine the requirement and role of ATR in cell proliferation. Using B cells as a model system to study proliferation, the authors conditionally delete ATR or the ATR kinase activity using stage-specific cre-deletor strains. They find that while loss of ATR is essential for early B cell development in the bone marrow, its loss in naïve B cells does not lead to an overt phenotype under homeostatic conditions. However, when naïve ATR-deficient B cells are activated in vitro, there is a severe defect in B cell proliferation. The authors then carry out a series of PI staining and BrdU-incorporation experiments to demonstrate that the proliferation defect is likely due to massive stalling of replication forks. They further demonstrate that ATR-deficient B cells can initiate DNA replication in early S but they fail to maintain the replicative state by mid-S phase due to reduced nucleotide levels and fork stalling. Finally, the authors show that productive DNA replication can be restored by inhibiting enzymes that suppress origin firing such as CDC7 and CDK1. Taken together, the authors suggest that in unstressed cells, ATR supports proliferation by modulating origin firing in early S-phase cells to avoid exhaustion of nucleotides during the later stages of S phase.
The role of ATR in the cellular DNA damage response following induced DNA damage has been extensively studied for the past several years. It is now clear that replication stress activates ATR and ATR-dependent phosphorylation elicits a concerted response that includes suppression of firing of replication origins. However, the potential role of ATR in regulating origin-firing in unstressed cells have not been fully investigated. In this regard, this study is a detailed characterization of B cells lacking ATR protein or without ATR-kinase activity. The experiments are well done and beyond the cell cycle/proliferation experiments that directly address the ATR-origin firing link, the data from the metabolomic, transcriptomic and the CRISPR/Cas9 screen will be an excellent resource for the field in general. However, the authors should address the issues below to strengthen their study.
1. Fig. 1. and S1: The authors have characterized CSR and the nature of the switch junctions in control and ATR-deleted B cells. The authors should test if the few cells that have switched have done so despite loss of ATR or if the switched cells have escaped cre-mediated deletion. 2. It would be informative if the authors could provide a comprehensive analysis of the B cell phenotype, including germinal center frequency at homeostasis (in Peyer's patches) and following immunization (with say NP-CGG), and the levels of serum Igs. 3. Fig. 2f. The authors claim that ATRi treatment at 46h post-stimulation has no effect on BrdU incorporation. However, the analysis is done 2hrs after ATRi treatment (at 48hs). How much difference is expected in this 2hr window in any case? It is very difficult to interpret this result without data showing how much BrdU normally gets incorporated between 46h and 48hrs. 4. The authors claim that even at 48h post-stimulation, ATR-deficient cells are viable. The authors should include some apoptotic markers to ensure that the cells have not primed to die even though they appear viable. 5. It would be informative if the authors could examine the nature of chromosomal abnormalities in metaphases of ATR-deficient cells at 48h post-stimulation. 6. The authors should be a little conservative in stating that they are testing the role of ATR in unstressed cells. The stimulated B cells are dividing rapidly and have started to express AID which has the potential to cause both Ig and genome-wide DNA breaks. Thus, to refer to ex vivo stimulated B cells as unstressed does not give a true picture of the cell type they are using.
Minor points. 1. The authors should describe the two ATR alleles in more details. They have been described earlier but without any description it was initially difficult to follow the text 2. In Fig. 1b, the authors should provide absolute cell numbers We thank all the reviewers for their encouragement and their thoughtful suggestions. The reviewers praised the importance and significance of studying normal ATR function in primary cells and the value of the definitive genetic approaches used to study an essential gene in our manuscript. The reviewers also pointed out some limitations of our study and made thoughtful and constructive suggestions. In the revised manuscript, we have included 16 new or updated figure panels and tables. They are Fig.1c, 1f, 1g, 3f; Sup. Fig. S1d, S1e, S1f, S1j, S2a, S2b, S2c, S2d, S2f, S2h, and S3d, and Sup. Table 1. We also made all the suggested text modifications and included additional references as suggested by the reviewers. We extended the discussion of our results in the context of prior studies using small chemical inhibitors and transformed human cancer cell lines. We believe that we have addressed all the major concerns of the reviewers. We sincerely appreciate the help and advice from the reviewers and the editors, which significantly improved our manuscript. Enclosed are the pointby-point responses to the review. The original reviewer comments are italicized and underlined here for easy identification. New and updated figures are also inserted into the point-by-point response (with yellow highlight) for convenience.

Reviewer #1 (Remarks to the Author):
The manuscript "ATR kinase supports normal proliferation in the early S phase by preventing replication resource exhaustion" by Menolfi et al. explores the role of ATR in DNA replication on unperturbed B cells. The authors take advantage of synchronous cell cycle entry of B cell upon cytokine stimulation to knockout ATR in non-dividing cells and observe the effects of the knockout on DNA replication upon stimulation. While the essential function of ATR in cell cycle has been extensively studied before, using ATR inhibitors and cancer cells, the authors of the current manuscript take it one step further, using primary cultures and a genetic approach. The main conclusion of the current study is that ATR is required to suppress excessive origin firing during DNA replication to prevent nucleotide depletion and subsequent DNA damage. This has been documented in cultured cancer cells before, so the main novelty is demonstrating that the same principles apply in primary B cells. The question why nucleotide complementation failed to restore S-phase progression remains unanswered. While the manuscript contains a vast amount of data and most of the conclusions are justified, the conclusions are mainly confirming the results that were previously reported in cancer cells/cell lines. In order to make it more novel, I propose either looking into some unsolved questions related to the effect of ATR inactivation (insufficiency of nucleotide complementation, effects on the nucleotide balance, etc.) or focusing on possible differences between healthy B cells and cancer.
We thank the reviewer for his/her encouragement and for highlighting the value of our study of primary cells and genetic systems. During the revision, we generated v-abl kinase transformed B cell lines carrying the same Atr +/C and Atr C/-alleles Ruzankina et al. 2007) and the tamoxifen (4OHT) inducible ER-Cre recombinase to determine the impact of ATR loss in transformed (in contrast to primary) B cells. Upon ATR deletion, there is a moderate loss of viability seen on Day 7 and a moderate accumulation of BrdU-negative S phase cells in the v-abl kinase transformed B cell lines (New Figure S1j). But overall, the loss of viability is significantly less than in primary B cells (from 85.8 to 81.8% in 4 days in transformed B cells vs. from 82.1% to 38.7% in primary cells, Figure S1c). This delay in cell death is not due to incomplete deletion of ATR, as confirmed by sensitivity PCR assay (New Figure S1k). These results highlight the crucial role of ATR kinase in primary cells that initiates the cell cycle from the G0 phase with low metabolic activity and limited reserve. In contrast, transformed cancer cell lines in continuous culture can benefit from resources diluted from prior cell cycles. Consistent with this model, when activated primary B cells were treated with ATR inhibitor at 24hr after activation (instead of New Figure S1j and S1k shows that v-abl kinase transformed B cell lines can tolerant ATR deletion (verified by PCR in k) much better than primary B cells (in Figure 2). We noted that the null allele (-) is at different exons from the conditional and del allele that is generated from LoxP recombination from the conditional allele. Therefore, the null allele appears like WT in the PCR designed to detect the C and del allele (in k). from the beginning), the disruption to DNA replication (measured by BrdU pulse chase) was less pronounced. See the updated Figure 3e red labeled, ATRi after 24hr vs. the blue labeled ATRi from time zero). We now include this in the text and discussion. We sincerely thank the reviewer for his/her insightful suggestions.
The question about the failure of nucleotide complementation is indeed exciting and challenging at the same time. Consistent with prior findings in cell lines ) and in yeast models (Forey et al. 2020), we were also NOT able to rescue the productive replication in ATR deficient cells with nucleoside supplementation alone. In addition to reporting and confirming this negative finding, we made three steps forward toward the mechanism. First, in addition to ribonucleoside (N) used in prior studies, we supplemented the cells with deoxy-nucleotide (dN) that bypasses the Ribonucleotide reductase (RNR). This approach is uniquely suitable for lymphocytes since lymphocytes have high expression and activity of deoxycytidine kinase (dCK) that can directly phosphorylate deoxynucleosides (dNs) to mono-phosphate nucleotides (dNMPs), while most other cell types can only phosphorylate nucleosides (ribo form) and then depend on RNR to convert NDP to dNDP (see diagram on the right). Indeed, mouse models lacking dCK display specific lymphocytopenia without affecting other major tissues (Toy et al. 2010). Still, dN cannot restore productive replication in ATR-deficient B cells, suggesting RNR is not the only rate-limiting factor. Second, with help from Dr. Baek Kim, we measured the intracellular dNTP levels ( Figure 6a) before and after dN supplementation and demonstrated that the dN supplementation successfully restored the dNTP levels in ATR-deficient cells. This has not been done in the previous studies and has left intake or conversion defects as possible reasons for the rescue failure. In this regard, we also showed that the dN supplementation significantly attenuated the DNA damage response ( (Figure 7g). To our knowledge, this is the first time the restoration of the dNTP levels in ATR-deficient cells has been documented along with productive replication. Collectively, these results support a new model in which the overuse of dNTP and the lack of other replication factors, including but not limited to dNTPs,causes replication failure in the ATR-deficient cells. This model explains why nucleotide supplementation has failed to rescue DNA replication without ATR. We thank the reviewer for his/her thoughtful suggestions, and have now added a paragraph to discuss this in the text. (Buisson et al, 2015). Here the authors conclude that ATR is mainly important for early S-phase cells based on the experiment where ATRi was added at different timepoints during cell cycle progression (figure 2f). On this figure the authors compare 2h ATRi incubation with 48h ATR incubation. Given that the cells enter S-phase less that 24h after activation, the 48h sample has been lacking ATR activity through the whole cell cycle, while 2h sample has been without ATR activity for only 2h. Similarly, on figure S3c, the authors compare 24h or 14h ATRi treatment to 2h. ATRi treatment in early S should be limited to 2h as well -my guess is it would not completely stop EdU incorporation.

ATR being most important in early S-phase was previously proposed by Zou lab
Indeed, we have thought about and tried these experiments suggested by the reviewer. The results are now included in the updated Figure 3e. Briefly, we stimulated the primary B cells and split the culture into four groups and added ATRi at 24hr after initial stimulation for ~1 hr (green line and green label) or for 24hr (red The diagram shows the simplified pathways from ribonucleoside (N) or deoxyribonucleoside (dN) to the dNTP. Lymphocytes have very active dCK that can effectively convert dN to dNMP, therefore bypassing RNR that was down regulated in ATRdeficient cells.
line and red label) and chased the BrdU+ cells (pulse-labeled for 30 min) for the next 24hr. As expected from the ATRi supplemented at 22h, the one hour treated and chase has no measurable impact on S phase progression. In contrast, when ATRi was added at the beginning (blue line), there was a significant block in the S phase progression. We also repeated the experiments independently a few times. The data is now included in the updated Figure 3e and quantified in New Figure S3d. Fig. 3e, where authors look at the ability of the cells to complete the cell cycle by quantifying the G1 phase cells (this could be repeated and statistics could be presented). However, here we can see that at 24h timepoint when the BrdU pulse is performed, much fewer cells in the ATRi samples are actually incorporating BrdU (13.5% vs. 36%), so this must be taken into account when quantifying the percent of BrdU+ G1 cells at 48h -the percent cells labeled with BrdU at 24h cells making it to G1 (and not the percent of all cells ending up BrdU+ in G1) may actually be higher in the samples with continuous 48h ATRi treatment (possible re-activation of CHK1 by DNAPK proposed by Buisson et al., 2015?). Additionally, the percent of BrdU+ cells appears to increase between 24h and 48h, which is difficult to explain given a short 30 min pulse.

A better experiment is presented on
We thank the reviewer for his/her thoughtful suggestion. We did have independent repeats and now also quantified them as a percentage of BrdU+ cells (control for BrdU labeling percentage difference). The results are now provided in updated Figure S3d. We also note that while ATRi treatment from time zero reduced BrdU+ cell percentage (potentially due the dominant negative role of the inactive ATR protein previously reported ), Atr KO B cells (CD21Cre+Atr C/-) had the same percentage of BrdU+ cells at 24h as the control Atr C/+ B cells (Updated Figure 3e and S3d), while the G1% among BrdU+ cells markedly reduced in Atr-KO cells. We thank the reviewer for noting the increased BrdU+% during chasing in some experiments. We only saw this in primary B cells and have tried multiple experimental conditions, which yielded variable results. One possibility might be the BrdU negative population that reflects non-responders to cytokine stimulation might have dropped out during the chases. As seen in Figure S1C, by day 4 the viability for AtrC/+ cell is only 75% and for AtrC/-cells is ~40%.
We also noted the results from the Buisson et al., 2015 paper. Indeed, we have measured pCHK1 at both 24hr ( Figure S3g) and 48hr (Fig. 2E) after activation. In neither case did we observe significant phosphorylation of CHK1. We noted that Buisson et al. performed their experiments in human cancer cell lines (Buisson et al. 2015). Here all our experiments were carried out in primary murine cells. We and others have previously shown that the protein levels and activities of DNA-PKcs and KU (the DNA binding component of DNA-PK) are Updated Figure 3e and S3d: 3e) Primary B cells were stimulated together and split in four different cultures. When ATRi was added at 24h post-stimulation, either for 1h (green treatment) or for the next 24h until cell collection at 48h (red treatment), BrdU incorporation and recovery resemble the one of the untreated ctrl. Comparable BrdU-positive G1 sub-populations were observed within the total of S phase cells, suggesting no defects in cell cycle progression and re-entry. However, when ATRi was added in G0, at the beginning of the experiment (blue treatment), cell cycle re-entry was severely compromised, revealing the essential function of ATR. The quantification of this data is now included in Figure 3e (shown as percentage of G1 cells among the BrdU+ cells as suggested by Reviewer 1). S3d) As suggested by the reviewer, we quantified the % of G1 BrdU+ cells versus total BrdU-labelled positive cells. CD21-Cre AtrC/-and ctrl cells treated with ATRi for the length of the experiment display a significant reduction of G1 BrdU+ cells, indicative of defects in cell cycle re-entry. Two or three independent experiments are quantified, and the graph refers to the representative flow cytometry analyses reported in Figure 3e. The quantification is reported in Supplementary Figure 3d.

S3d
Page 4 of 16 50-100 fold lower in mouse cells than in human cells (Jiang et al. 2019). This might explain the lack of DNA-PKmediated CHK1 phosphorylation in our murine-based experiments. We now mention this in the text. We are very interested in this species-specific difference but think that this is beyond the scope of the current manuscript.

Metabolome profile on figure 5d is remarkable. It is showing lower concentrations of the majority of nucleotides in ATR-deficient cells, however, ATP, GTP, UTP, CTP, and dGTP appear to have increased with
the loss of ATR, but it not discussed at all. These data indicate specific biosynthetic pathways may be regulated by ATR. A recent preprint from Bakkenist lab (Sugitani et al., 2022) showed regulation of RRM2 by ATR in dividing primary T cells; could a similar mechanism explain this phenotype in B cells?
We thank the reviewer for pointing out the increase in NTPs. We also noted it in the main text. While we do not know the exact cause of the moderate NTP increase (1.2-1.5 fold), it would be consistent with the lack of rescue by nucleosides (ribo form) reported previously . It is possible that the lack of NTP usage in some pathways contributed to this accumulation. Alternatively, there might be excess conversion to NTP from NMP and NDP. As reported by Bakkenist and colleagues (Sugitani et al. 2022), we also observed a 1.2-1.5 fold reduction of the RRM1 and RRM2 mRNA levels in the Atr-deficient B cells ( Figure  S5e). This and other observations encouraged us to supply the cells with deoxynucleosides (dN) rather than nucleosides to bypass RNR (see the response to reviewer 1 general comments and diagram) (Lane and Fan 2015). While dN successfully increases deoxy nucleotide levels in the cells, it could not rescue productive DNA replication (measured by BrdU incorporation) ( Figure 6). (Petermann et al. 2010) with CHK1 inhibitors, the role of CDK1 in ATRi-induced effects has been described by Bakkenist lab .

Throughout the manuscript, the authors compare their findings in B cells to the studies in yeast, yet they fail to reference some key studies done in mammalian cells -nucleotide complementation has been done with CHK1 depletion by Gottifredi lab (González Besteiro et al, 2019); CDC7 inhibition has been tried in human cells by Helleday lab
We thank the reviewer for pointing out those key references that we missed. We have now cited them all in the main text-González -Ref 38, Petermann et al., 2010and Moiseeva et al., 2019-Ref 62. We further note that CHK1 and ATR have independent functions beyond their well-recognized epistatic roles. In contrast to CHK1, which is transcriptionally silenced and absent in G0 quiescent cells ( Figure S3g), ATR is transcribed in G0 arrested primary cells (Lee et al. 2014a). Moreover, the three references all use cancer cell lines treated with either small molecule inhibitors for ATR or CHK1, highlighting the knowledge gap in non-transformed and genetically deleted cells. Together with prior findings, our data collected on primary B cells show that the ATR kinase activity suppressing the excessive firing of the replication origins to coordinate S phase progression is a conserved and essential function of ATR in both normal and transformed cells. between each paired IdU fiber and considered ratio =1 as a symmetric fork and ratio >1 as fork asymmetry. The ratio was computed using the longer IdU fiber as the numerator and the shorter as the denominator. The data indicate that similar to what has been previously reported for CHK1 inhibitor-treated cells, replication forks are asymmetric in CD21-Cre + Atr C/-cells compared to ctrl cells. We now report this in new Figure S2d. We have re-plotted the BrdU/PI plots (Updated Figure 2a and 2f, as well as Figure S2a and S2b) to make them vertical and consistent with the standard view in the field in all main and supplementary figures.

Reviewer #2 (Remarks to the Author):
The study by Menolfi et al. entitled "ATR kinase supports normal proliferation in the early S phase by preventing replication resource exhaustion" aims at understanding the role of ATR in the proliferation of unstressed proliferative B-cells. They initially show that the genetic deletion of ATR, or the loss of its kinase activity, impairs clonal expansion in-vivo, but has no effect on G0-arrested naïve

B-cells. To better understand the molecular mechanism by which ATR promotes B cells proliferation, they use a model of naïve B cells ex-vivo culture, stimulated by cytokines (mimicking B-cells activation) and used stateof-the-art techniques such as metabolomics and CRISPR-Cas9 screens, to uncover the determinants of ATRdependent proliferation. The authors show that cells depleted of ATR, show defects in S-phase progression, and propose that this is due to the function of ATR in fine-tuning origin firing to limit dormant origin firing and prevent a shortage of replication factors, and/or dNTPs. The manuscript tackles an appealing aspect of ATR functions in normal proliferation and uses a relevant physiological model of B-cell activation. These observations are potentially interesting, but the novelty is limited by the fact that the role of ATR and CHK1 in controlling origin firing locally and globally, has already been well established in the context of replicative stress. The study suffers a lack of evidence supporting their views and few technical limitations. Therefore, I do not think that this manuscript is suitable for publication in Nature Communication in its current form.
We thank the reviewer for carefully elevating our manuscript and for his/her candid feedback. We note that ATR (and CHK1 and other proteins) has many functions during replication stress. But not all of these functions are essential for normal unstressed primary B cells. While it is important to understand the role of ATR under replication stress or in cancer cells, the success of ATR inhibition as a therapeutic approach also requires a thoughtful evaluation of ATR function in normal unstressed tissues and cells in vivo. To some degree, it is satisfying and comforting to know that primary cells did NOT re-invent the wheel. Instead, the conserved function of ATR in suppressing excessive origin firing (among many other functions of ATR) underlies the essential role of ATR in proliferation. With this knowledge, it would be possible to target other cancer-specific and potentially replication stress-specific functions of ATR for cancer therapy while sparing normal tissue/cells.

1-The authors show that CDK1/CDC7 dual inhibition restores the proliferation of ATR-deficient cells. This is a critical result that has potential applications in several pathologies. However, the relevance of these findings needs to be confirmed in an in-vivo setting, such as in the context of naïve B cell clonal expansion
We agree with the reviewer. To determine the in vivo implication of Atr-deficiency in B clonal expansion, we adopted a well-established immunization model (Klein et al. 2003) to examine the impact of Atr-deficiency in naïve B cell germinal center response after antigen exposure (e.g., sheep red blood cells-SRBC). In CD21Cre + Atr C/-mice, both spontaneous and induced germinal center B cells (GL7+ and CD95+) -the fraction of B cells activated by T cells upon antigen exposure in vivo -were significantly reduced (New Figure 1f and  1g), supporting a critical role of ATR in B cell clonal expansion. While there are many clinical-grade CDK inhibitors, most target CDK4/6 or CDK12/13. Unfortunately, while the current CDC7 inhibitor we used is in a phase I clinical trial, the CDK1 inhibitor is unsuitable for in vivo application, preventing us from rescuing the germinal center response in vivo.

2-Linked to major point 1, do patients with Seckel syndrome suffer from a deficit of naïve/activated B-cells? Is it recapitulated in the mouse model of Seckel syndrome?
We do not have direct access to Seckel syndrome patients. On the OMIM database maintained by NCBI, there are 5 ATR mutation alleles from 4 different case reports of Seckel syndrome patients. They all have variable levels of ATR, consistent with hypomorphic ATR deficiency (https://omim.org/entry/601215#allelicVariants). All the patients display dwarfism and microcephaly. The blood cell counts, and more specifically lymphocyte counts, were not discussed in any of the case reports. However, in one study, an EBV-transformed lymphoblastic cell line was derived from a patient carrying compound ATR-heterozygous mutations. The authors comment in the text that the cell line grows slower (Mokrani-Benhelli et al. 2013), consistent with the role of ATR in B cell proliferation.
Meanwhile, the hypomorphic Seckel syndrome mouse model (Murga et al. 2009) does show reduced white blood cell counts ( Figure 3e). Moreover, using bone marrow transplantation, the authors showed that AtrS/S bone has a significant yet reduced ability to reconstitute the B and T cell linage specifically (in their Sup. Figure S6e). We note that the authors suggested that hematopoietic niche defects might contribute to the pancytopenia in the Seckel mouse models, further highlighting the value of using lineage-specific and developmental-stage specific Cre alleles to dissect the cell-autonomous and development stage-specific role of ATR in B lymphocytes presented in our study.
New Figure 1f (left) and 1g(right -bar graph) show the frequency of activated germinal center B cells (marked by GL7+ and CD95+) among all splenic B cells has significantly reduced both at the basal line (naïve) and upon immunization (sheep red blood cells). The IHC staining for the germinal center marker Bcl6 validated the flow cytometry findings. On the right, the number of germinal center B cells after immunization was plotted from 4 independent mice of each genotype (average and standard errors were plotted). The student t-test was used to determine the p-value. Finally, we note that lymphocytes are not essential for viability or embryonic development. Given the pleiotropic role of ATR in nearly all cell types, it is not surprising that lymphocytes are not the primary cause of morbidity in ATR-mutated patients. This is different in adult animals, where most tissues have stopped proliferation and are insensitive to ATR loss. At the same time, lymphocytes, upon activation, undergo rapid expansion, highlighting the importance and the unique value of B cells-specific conditional ATR deficient alleles. Moreover, we note that B cell development requires rapidly expanding large Pre-B cells after successful IgH heavy chain rearrangement. If ATR is essential for the proliferation of primary cells, the reduced B cell numbers in the Seckel Syndrome model are likely due to development defects (not necessarily naïve/activated B cell defects). The activation defects are crucial for adult patients exposed to ATR inhibition as part of cancer therapy. Here we conditionally deleted ATR only in naïve/mature B cells (after the expansion of pre-B cells), bypassing the early need for ATR in B cell development, and allowing us to directly access the response to ATR loss in naïve primary B cells.

3-The authors should show representative images of the DNA fibers, and estimate the inter-origin distance, or the percentage of "first-labels origins" to support their findings on origin firing.
As suggested by the reviewer, we have included examples of DNA fibers in New Supplementary Figure 2c and 2d. Unfortunately, we cannot access the instrument to perform the inter-origin distance measurement in our lab right now. Given our focus on primary B cells freshly isolated from the animal and activated within 24hrs, it would also be challenging to ship the cells to outside collaborators for this experiment. We have discussed these caveats in our paper and cited others' publications in which inter-origin distance has been measured in the context of ATR inhibition in murine and human cells Sugitani et al. 2022).

4-Analysis of cell viability by SSC/FSC gating is not accurate and can be misleading. The authors should confirm those results with a viability dye.
Following this reviewer's comment, we compare the SSC/FSC gated live population with the standard PI-Annexin staining in murine B lymphocytes. As seen in this figure for review only, in murine B lymphocytes, parallel analyses for SSC/FSC and PI-Annexin show that SSC/FCS has a ~98% accuracy in detecting apoptotic cells (the sum of early and late apoptosis   SSC/FSC to gate out live vs. dead cells is routine. This super ability for SSC/FSC to distinguish live vs. death in lymphocytes is partly due to the nearly perfect sphere shape and small cytoplasm of lymphocytes, which give little noise in SSC measurements.
Nevertheless, we activated B cells from three or more independent CD21Cre+Atr+/C and CD21Cre+AtrC/mice and performed apoptosis analysis using Annexin and PI. The result is presented in New Figures S1e and 1f and shows that, indeed the activated ATR-deficient lymphocytes undergo apoptosis.

5-In Supplementary Figure 2d, Hydroxyurea did not affect S-phase progression? Was it used at a suboptimal concentration? The authors should confirm that HU and the ATM/DANPKcsi inhibitors hit their targets in this cellular model.
We thank the reviewer for this comment. Original Supplementary Figure 2d is now Figure S2g. We indeed used a low dose of HU (20 µM instead of 2mM) to allow the cells to successfully enter S phase. Otherwise, we would not be able to measure BrdU levels in cells with S phase DNA content (the goal of this experiment). We also performed experiments with a high dose of HU 2 mM, which prevented S phase entry as shown in New Supplementary Figure 2h. Therefore, HU did work appropriately. We have used commercial (from Selleckman) ATMi, and DNA-PKi at the dose indicated (7.5-15uM) (Crowe et al. 2018;Jiang et al. 2019;Crowe et al. 2020;Shao et al. 2020;Wang et al. 2020) and always confirmed that ATMi reduced Ig class switch recombination (CSR) by ~50% (Lumsden et al. 2004;Reina-San-Martin et al. 2004;Pan-Hammarstrom et al. 2006) as also reported in the figure for reviewers only. In comparison, DNA-PKcs inhibitor reduced CSR by ~10-20%, consistent with DNA-PKcs null cells (Manis et al. 2002;Crowe et al. 2018;Crowe et al. 2020). Given the amount of data and figures in the current manuscript, we felt that it is not necessary to include these technical validations of commercially available and well-characterized inhibitors in the main data. We and others have also reported that ATM null (Lumsden et al. 2004;Reina-San-Martin et al. 2004;Pan-Hammarstrom et al. 2006) or DNA-PKcs null (Manis et al. 2002;Crowe et al. 2018;Crowe et al. 2020) B cells show no proliferation defects. We have now cited those papers in the revised manuscript.
New Figure S1e and S1f :Representative Flow cytometry for apoptotic markers (PI and Annexin) in activated lymphocytes (S1e) and quantification (S1f) for different days after stimulation. The data proved that the Atrdeficient B cells indeed died of apoptosis and demonstrate excellent ability for FSC/SSC to separate the live vs dead lymphocytes.
New Figure S2h :High dose HU completely blocked DNA replication. To quantify the frequency of BrdU negative S phase cells, we have to move to a lower dose of HU.

Figure for reviewers.
Upon activation, CSR cells were either left untreated or treated with ATMi (7.5 µM). Cells were collected after 4 days in CSR medium and analyzed by flow cytometry for IgG1 and B220. ATMi treatment led to ~50% reduction in CSR efficiency. Figure 2E, the authors should show the expression of ATR by Western-Blot.

6-
Unfortunately, there is no reliable antibody for mouse ATR. The original antibody used by Dr. Eric Brown in his first publication on ATR KO mice is not available anymore. Despite several efforts, including reaching out and testing older stocks provided by Dr. Brown's lab, we failed to detect mouse ATR. Meanwhile, we note that this ATR conditional allele has been very well characterized since 2000 (Brown and Baltimore 2000;Brown and Baltimore 2003;Ruzankina et al. 2009;Onksen et al. 2011), and we have documented robust deletion in primary B cells via PCR ( Figure S1b and S1k). Perhaps most importantly, deletion of ATR in pre-B cells via Mb1Cre completely abrogated B cell development in vivo, and deletion of ATR in naïve B cells via CD21Cre blocked B cell activation within one cell cycle and erased CHK1 phosphorylation (Figure 1 and Figure 2). Altogether, these solid and undeniable phenotypes, together with a sensitive PCR assay for ATR deletion, indicate that ATR is indeed deleted in naïve B cells. We noted that naïve B cells could stay in the peripheral organs for months, if not years, which would be more than sufficient to overcome the long half-life of ATR proteins. The raw data and the adjusted p-values for consistent hits has been included in Supplementary table 1, now with updated highlights. Moreover, the entire CRISPR dataset has been made available at GSE214643.We opted to use FDR< 0.2 because, 1) several recent CRISPR Screens published on DNA repair and genomic instability use this standard (Hart et al. 2017;Noordermeer et al. 2018;Zimmermann et al. 2018;Olivieri et al. 2020) and 2) the developer of the MAGeCK pipeline (https://sourceforge.net/p/mageck/wiki/Home/) that we used to analyze the CRISPR/Cas9 screen suggests this FDR for identifying recurrent targets in multiple independent screens (3 ATR inhibitor and 1 CHK1 inhibitor screen here). The detailed date including adjusted FDR, Z and beta scores of the hits are included in updated Sup. Table 1.

8-
The findings from the screens are very interesting but I see a biological bias that the authors did not comment on. How many of these gRNAs totally prevent origin firing? In this situation, they would confer resistance to ATRi, not because they are ATR targets but because they impede S-phase entry. For example, I would expect to find gRNAs of genes that induce G1-arrest (Mdm2, p21 ubiquitin ligases,…).
We thank the reviewer for this comment. We do see a few genes associated with G1 cell cycles (e.g., Cyclin D3 in individual screens). We do not expect to see targets that "completely" block origin firing. CRISPR screening is an enrichment screen based on cell proliferation ability. If the cells are completely arrested or die, we would not see any enrichment. Similarly, our rescue with CDC7i and CDK1i used a sub-lethal dose. A high dose of CDC7i and CDK1i would completely block WT B cells. In fact, we chose the dose of CDC7 and CDK1 inhibitors that do NOT affect the proliferation of control B cells. Finally, we noted that primary B cells have a different p53 response than common epithelial cells. Specifically, upon radiation-induced p53 activation, primary lymphocytes activate apoptosis rather than cell cycle arrest. For this reason we also did not observe p21 itself as a reliable p53 target in primary lymphocytes.
Minor points 1-The authors did not mention the gating strategies used in figure 1a before the B220/CD43 and B220/IgM plots. Did they exclude dead cells (e.g. 7AAC-or DAPI-cells)?
The dead cells were excluded using FSC/SSC, and red blood cells were excluded using a TER119 antibody. We now discussed this in the methods. As shown in response to reviewer 2 major comment 4, in lymphocytes, SSC and FSC are reasonably good indicators for viability. That information is now included in the online method section.
Page 10 of 16 2-The statistical analysis is missing in figure 1b.
We now included the statistical analysis for B cell populations and also absolute bone marrow cell counts (updated Figure 1b and 1c). Figure 7A, the legend is missing Figure 7A legend has been expanded to include more details now.

4-the authors show that ATR depletion induces DNA damage in quiescent B-cells. This is rather surprising. Can the authors comment?
Indeed, we were very interested in this observation. Literature suggests that ATR might participate in nucleotide excision repair (Lee et al. 2014b) and be activated by transcription-generated single-strand RNA (Gorthi et al. 2018). In Figure S3h, we showed that despite the increased alkaline comet signals, we did not find significant phosphorylation of KAP1 or RPA, likely consistent with low levels of single-strand nicks. We feel that this discussion is beyond the scope of this already very complicated paper, but interested in pursuing this question in future studies.

Reviewer #3 (Remarks to the Author):
In this manuscript, Zha and colleagues examine the requirement and role of ATR in cell proliferation. Using B cells as a model system to study proliferation, the authors conditionally delete ATR or the ATR kinase activity using stage-specific cre-deletor strains. excellent resource for the field in general. However, the authors should address the issues below to strengthen their study.
We thank this reviewer for his/her encouragements. We have addressed the specific comments below.

Fig. and S1: The authors have characterized CSR and the nature of the switch junctions in control and ATR-deleted B cells. The authors should test if the few cells that have switched have done so despite the loss of ATR or if the switched cells have escaped cre-mediated deletion.
We thank the reviewer for his/her thoughtful suggestions. Indeed, we performed PCR genotyping on different days after CSR routinely. There is a variable amount of residual ATR conditional allele that often flares up in D3 or D4 only in the Atr C/-mice. This is consistent with the strong selection against ATR deleted cells and the accumulation of the escapers. But as seen in the representative figure, the deletion band is always the dominant band in the PCR, suggesting the majority of the cells have deleted ATR. A similar observation has been made by us and others on CtIP, another essential gene in activating B cells (Polato et al. 2014;Liu et al. 2019;Wang et al. 2020). We have now included one representative data here for the review.

It would be informative if the authors could provide a comprehensive analysis of the B cell phenotype, including germinal center frequency at homeostasis (in Peyer's patches) and following immunization (with, say, NP-CGG) and the levels of serum Igs.
Due to the small size of our animal colony after COVID related shutdown (we lost 80% of our mice in 3 days), we have only a limited batch and ctrl mice for these experiments right now. Instead of the antigen-specific immunization (NP-CGG), which usually yields small germinal centers and viable antibody titers, we chose to immunize the mice with sheep-red-blood cells that generate a robust and reliable germinal center response (new Figure 1f and 1g). Indeed, the frequency of GC B cells (CD95+ and GL7+) is reduced in CD21Cre+AtrC/mice. Moreover, upon immunization, the induced GC cell frequency is also lower in CD21Cre+AtrC/-mice. Together the data provide strong evidence for the critical role of ATR in germinal center response. Due to the number of animals needed and the duration required for multiple rounds of immunization to measure antigenspecific Ig titers, we were not able to measure antigen-specific serum Ig levels in vivo. But we felt that the additional evidence on both Germinal center flow cytometry and histology analyses before and after immunization uniformly point to a B cell maturation defect associated with ATR loss.

PCR Genotyping during activation for the Review.
Here we show in the course of the 4 day stimulation, the conditional bands remain at very low levels if any. The rather prominent aspecific band on D0 might reflect impurity during magnetic beads purification, which usually achieves 90-95% B220+ cells. During the course of stimulation, the deletion is best at D1 and D2, and in the AtrC/-B cells, after D4, there are very few viable cells left, rendering the PCR unreliable.
Page 12 of 16 3. Fig. 2f. The authors claim that ATRi treatment at 46h post-stimulation has no effect on BrdU incorporation. However, the analysis is done 2hrs after ATRi treatment (at 48hs). How much difference is expected in this 2hr window in any case? It is very difficult to interpret this result without data showing how much BrdU normally gets incorporated between 46h and 48hrs.
When performing BrdU labeling, all the cells that were in the S phase during the labeling window will be BrdU positive. Similarly, when we treated the cells with ATR inhibitor for 2 hrs, all the cells, including all S phase cells, will lose significant ATR activity in this window. The point of this updated Figure 2f is to prove that ATR is not essential for the ongoing S phase and on-going DNA replication; as such, inhibiting ATR does NOT immediately stop BrdU incorporation as high dose HU would do (New Figure S1h). We now better explain this in the text.
We also treated the cells for 24h (from 24hr to 48hr after activation) and chased the BrdU+ cells (S phase cells during the BrdU pulse label). Updated Figure 3e and New Figure S3d (quantification) -see the response to review 1 comment 1 -now show that once the cells have started proliferation, ATR inhibition has less immediate impacts.
4. The authors claim that even at 48h post-stimulation, ATR-deficient cells are viable. The authors should include some apoptotic markers to ensure that the cells have not primed to die even though they appear viable.
We have now performed Annexin-PI staining for the activated B cells (Control and Atr deficient) (shown in new Figure S1e and S1f (see response to reviewer 2 major comment 4). There is significant cell death, so the analyses focus on the 50% viable cells. We noted that viability measured by FSC/SSC nicely aligns with those measured by Annexin-PI. The viable cells defined by FSC/SSC are 98+% viable using Annexin V and PI as the marker. (see response to reviewer 2 major comment 4).

It would be informative if the authors could examine the nature of chromosomal abnormalities in metaphases of ATR-deficient cells at 48h post-stimulation.
We thank the reviewer for his/her suggestion. We have performed Telomere-FISH in CD21-Cre Atr+/C, AtrC/and AtrC/KD activated B cells at D3 post-stimulation. We chose to use T-FISH to enhance the sensitivity to New Figure 1f (left) and 1g(right -bar graph) show that the frequency of activated germinal center B cells (marked by GL7+ and CD95+) among all splenic B cells have significantly reduced both at the basal line (naïve) and upon immunization (sheep red blood cells). The IHC staining for the germinal center marker Bcl6 validated the flow cytometry findings. On the right, the number germinal center B cells after immunization were plotted from 4 independent mice of each genotype (average and standard errors were plotted). The student t-test was used to determine the pvalue.
detect chromosome breaks, since mouse chromosomes have short arms, and loss of chromosome fragments can easily be missed without telomere markers (Franco et al. 2006). ATR-deficient B cells show a small but consistent increase in aberrations per metaphase, in particular, breaks and fragile telomeres. Given the severe cell cycle arrest, the very low level of mitotic cells, and the fact that the chromosomal abnormality was measured in metaphase, we believe that this likely under-estimates the actual number of breaks and instability in the S phase or G2 phase cells. Nevertheless, we have included the data in New Figure S2f and discussed them in the text. We thank the reviewer for this insightful comment. Indeed, AID expression might add another layer of instability. Prompted by this suggestion, we searched for when AID is expressed in activated B cells. We found the following references that indicate that AID protein is not detected in the first 24 hours after activation (Schrader et al. 2005;McBride et al. 2006). We have noted this in the paper and focused most data, including metabolome, RNA-seq, and dNTP data, on the first 24 hours after stimulation.
Minor points. 1. The authors should describe the two ATR alleles in more details. They have been described earlier but without any description it was initially difficult to follow the text We thank the reviewer for pointing this out. We have now included a detailed discussion about the alleles in the online method section. The Atr KO allele made by Dr. Eric Brown in the Baltimore lab replaced exon 1-3 of Atr with a Neo-R cassette. The ATR conditional allele and the corresponding Del () allele (after loxP recombination) introduced two LoxP sites flanking the C-terminal exons encoding the kinase domain. Since the Atr C/ and Atr KO involve distinct exons of the Atr gene, the PCR genotyping designed for Atr C/ would show a "WT" band in Atr KO cells. Similarly, PCR genotyping designed for Atr KO would detect the intact exon 1-3 in the New Figure S2f Top panel shows the frequency of chromosomal breaks identified by Telomere FISH. Lower panel shows the level of telomere instabilitydefined by more than 1 telomere signal per chromosome end.
Atr C/ mice. This can be seen in New Supplementary Figure 1k where the v-abl kinase transformed B cell line 8412 is Atr C/KO . In both cases, the ATR protein cannot be detected, resulting in a complete loss of ATR. In our studies, we used the Atr KO and Atr  alleles interchangeably and referred to them as Atrtogether for simplicity.

In Fig. 1b, the authors should provide absolute cell numbers
We have now included the absolute B cell number from one femur per mice in new Figure 1c for CD21Cre AtrC/-and C/+ mice (see page 9 of the point-by-point response). During COVID, we were asked to euthanize 80% of our colony in 3 days. Since we have collected all the Mb1Cre+Atr C/-data before COVID and knew that there are no naïve B cells in the Mb1Cre+Atr C/-mice for further study, we have not re-established the experimental Mb1Cre+Atr C/-colony. As a result, we currently do not have live animals at the proper age for cell count analyses. But since Mb1 is a B cell-specific Cre (Hobeika et al. 2006) and does not affect T cells and other lineages, the percentage of B cells in bone marrow provides an objective estimation of the B cell defects in those mice. We apologize for the lack of those data. We believe it would not jeopardize the overall conclusion, supported by Figures 1a, 1b.
In response to my comment #1 about various treatment times on figure 2f, the authors refer to a 1h treatment in the middle of S-phase on figure 3e that, as expected, had no effect on the ability of the cells to complete the cell cycle. However, my comment stated "ATRi treatment in early S should be limited to 2h", it was with regards to figure 2f, and this was not done. Figure 2f still compares 2h treatment to 48h treatment, and this is not very informative. If it is impossible to pinpoint the 2h of early S-phase, comparing the exposure for the first 24h only to the exposure for the second 24h (24-48) could be an option. A similar point was brought up by Reviewer #3, where the authors respond that "once the cells have started proliferation, ATR inhibition has less immediate impacts". The effects of ATR inhibition on proliferating S-phase cells have been studied repeatedly and it is clear that ATRi does not act like HU. What needs clarification is how "immediate" the effects on the early S-phase cells are. Because most of the treatments supporting this statement are 48h, which is not a shortterm treatment. How do the authors discriminate between the effects of the exposure length vs the timing of the exposure to a particular cell cycle phase? I think this is a key point to support the conclusion about the role of ATR specifically in early S-phase.
In response to my comment #4, the authors added three relevant citations, although they picked a wrong citation from Bakkenist lab -the WEE1 inhibitor study instead of the ATR/CHK1 study. It is worth noting that two out of these three articles use siRNA-mediated CHK1 depletion rather than kinase inhibition, and the third one includes data on the non-cancer BJ-hTERT cells, contrary to what the authors state in the rebuttal ("the three references all use cancer cell lines treated with either small molecule inhibitors for ATR or CHK1") and in the main article text ("Indeed, several previous studies have tried to counteract the toxicity of ATR or CHK1 inhibitors by inhibiting replication origin firing with variable degrees of success in human cancer cell lines"). The three articles also focus on the effects of ATR/CHK1 depletion/inactivation on origin firing and fork speed, and hardly mention the toxicity of the inhibitors. Therefore, my comment about properly discussing relevant literature is not fully addressed.
In response to comment #6 by Reviewer #2, the authors claim there is no good antibody for mouse ATR, but there is definitely at least one antibody that has been repeatedly used to detect mouse ATR by western blot -cell signaling #2790. Multiple citations are listed on the manufacturer's website, including three published articles that successfully used it to detect mouse ATR by western blot.